CHAPTER 3

Hydration Dynamics at Fluorinated Protein Surfaces

This chapter first appeared as an article in Proceedings of the National Academy of

Sciences, USA: Oh-Hoon Kwon, Tae Hyeon Yoo, Christina M. Othon, James A. Van

Deventer, David A. Tirrell, and Ahmed H. Zewail. Proc. Natl. Acad. Sci. USA 2010, 107,

(40) 17101-17106.

156

Abstract

Water-protein interactions dictate many processes crucial to protein function

including folding, dynamics, interactions with other biomolecules, and enzymatic catalysis.

Here we examine the effect of surface fluorination on water-protein interactions.

Modification of designed coiled-coil proteins by incorporation of 5,5,5-trifluoroleucine or

(4S)-2-amino-4-methylhexanoic acid enables systematic examination of the effects of side-

chain volume and fluorination on solvation dynamics. Using ultrafast fluorescence

spectroscopy, we find that fluorinated side chains exert electrostatic drag on neighboring

water molecules, slowing water motion at the protein surface.

Introduction

The past decade has witnessed substantial expansion in the number and diversity of

noncanonical amino acids that can be incorporated into recombinant proteins expressed

in bacterial cells (1-3). Fluorinated amino acids have drawn special attention (4-16)

because of the unusual solubility properties of fluorinated hydrocarbons. Several

independent studies have shown that fluorination of coiled-coil and helix-bundle proteins

leads to enhanced stability with respect to thermal or chemical denaturation (6-12), an

effect attributed to the hyperhydrophobic and fluorophilic character of fluorinated amino

acid side chains.

Although both classes of compounds are hydrophobic, hydrocarbons and

fluorocarbons differ in important ways (17-22). The high electronegativity of fluorine

renders the C-F bond both strongly polar and weakly polarizable (17, 21, 22). The dipole

associated with the C-F bond exerts strong inductive effects on neighboring bonds (23)

157 and can form reasonably strong electrostatic interactions with ionic or polar groups when

the two moieties are appropriately positioned. The hydrophobic character of fluorinated

compounds has been described as “polar hydrophobicity (17),” and is believed to play

important roles in organic and medicinal chemistry. Furthermore, the C-F bond is

significantly longer than the C-H bond, and the calculated volume of the trifluoromethyl

group is about twice that of a methyl group (20). The studies described here constitute an

attempt to understand more fully the interaction of water with fluorinated molecular

surfaces, and to provide a sound basis for the use of fluorinated amino acids in the

engineering of proteins with unique and useful physical properties.

The hydration layer adjacent to protein surfaces exhibits properties different from

those of bulk water; the more rigid and denser structure of the hydration layer plays a

crucial role in protein structure, folding, dynamics, and function (24-26). Elucidation of

the dynamic features of this region, on the timescales of atomic and molecular motion, is

essential in understanding protein hydration. In the past decade, the knowledge of

hydration on protein surfaces has been extensively expanded by studying the dynamic

properties of biological water for various proteins containing tryptophan (Trp) or

synthetic fluorescent amino acids as local probes; the results have revealed

multicomponent relaxation dynamics spanning a wide range of timescales (25, 27-29).

The nature of the protein hydration layer can be affected not only by the topographic and

electrostatic properties of the protein surface (24), but also by the physical and chemical

properties of individual surface-exposed residues (27, 30). In view of the unique

properties of the C-F bond and of fluorocarbon–water interfaces (23, 31), we anticipated

that fluorinated amino acid side chains might exhibit unusual hydration behavior. Here

158 we report studies of local hydration dynamics at fluorinated protein surfaces by

monitoring the time-dependent fluorescence Stokes shifts of surface-exposed Trp

residues in coiled-coil proteins with 5,5,5-trifluoroleucine (Tfl, 1; scheme 3.1) residues

adjacent to the probe. The results are compared to the hydration dynamics at

hydrogenated protein surfaces with Leu (2) or (4S)-2-amino-4-methylhexanoic acid

(homoisoleucine, Hil, 3) adjacent to the Trp probe. Hil has approximately the same

volume as Tfl (20, 21), and although the shapes of the residues differ, the nearly identical

side-chain volumes of Tfl and Hil allow us to differentiate changes due to fluorination

from those that result from the increase in side-chain volume that accompanies

replacement of Leu (scheme 3.1).

Results

Coiled-coil protein system. The coiled-coil protein A1 (figure 3.1 A and B) was used as a

model system to examine the effects of fluorinated amino acids on local hydration

dynamics. The primary structure of A1 contains six copies of a heptad repeat (abcdefg)n,

where positions a and d are occupied by hydrophobic amino acids. Self-association of the

peptide juxtaposes the a and d positions and results in the formation of a hydrophobic core.

Fluorinated Leu analogues have previously been incorporated into the d positions of A1;

the resulting proteins exhibited improved resistance to thermal and chemical denaturation

with minimal differences in secondary structure (9, 11, 12). In this work, the surface-

exposed Asp residue at the f position of the third heptad (position 34) was replaced by Trp,

which serves as a fluorescence probe (figure 3.1C). The Trp variant of A1 was designated

A1m. In order to examine the effects of fluorinated analogues on the local hydration

159 dynamics, a Leu codon was introduced at one of two positions within A1m. Mutation of a

serine residue at the c position of the third heptad (position 31) yielded a variant of A1

designated S31L (figure 3.1D), while replacement of an alanine residue at position b of the

fourth heptad (position 37) gave the A1 variant A37L (figure 3.1E). Each protein was

expressed in Tfl, Leu, and Hil form, yielding a total of nine different proteins that were

examined in detail (see (32) for nomenclature).

Characterization of global structure. Analysis of each protein showed that the overall

structural properties of the molecules were generally insensitive to genetic mutations and

incorporation of noncanonical amino acids. Circular dichroism spectroscopy indicated

that all nine proteins were helical, as determined from the molar ellipticity at 222

nanometers (figure 3.2) (33); an analysis with K2D2 software showed that the helicities

of individual proteins range between approximately 40% and 48% (34). These results are

consistent with the design of the A1 protein (35), in which approximately half of the

amino acids are located within the heptad repeats expected to form α-helical secondary

structure. The oligomerization states of the protein samples were determined by

sedimentation velocity analysis (figure 3.3). Although A1 forms dimers and tetramers at

neutral pH (11), the variants examined in this study form trimers or hexamers under

mildly acidic conditions (pH 4). We suggest that protonation of Glu side chains at the e

and g positions (figure 3.1A) of the proteins decreases the density of negative charges

adjacent to the hydrophobic core and promotes formation of larger helical aggregates at

pH 4. A1m, in which the single Trp residue occupies a surface-exposed position, is

predominantly trimeric in Leu-, Tfl-, and Hil-forms, with a small fraction of hexamers

160 (see figure 3.3). The majority of the S31L samples are present as hexamers, while the

A37L samples appear to contain mixtures of trimers and hexamers.

Characterization of local structure. The steady-state fluorescence emission spectrum of

Trp depends on the extent of exposure of the Trp side chain to water (36). All nine

protein samples showed emission maxima between 349 and 352 nm, close to that of free

Trp at 353 nm (table 3.1 and figure 3.4). These observations indicate that the Trp residues

are exposed to the aqueous environment (consistent with the original design), and not

involved in oligomerization of the proteins. In addition, the steady-state UV-visible

absorption and steady-state fluorescence emission spectra of each mutant containing Leu

were nearly identical to the spectra of the corresponding mutant when it contained Tfl or

Hil (figure 3.4), further confirming that perturbation of the protein structure upon

replacement of Leu by Tfl or Hil was minimal.

The mobility of the probe residue was explored in each protein by measuring fs-

resolved depolarization dynamics (figure 3.5). The anisotropic dynamics were found to

consist of three components: ultrafast (≤ 500 fs), intermediate (20–80 ps), and slow (≥2

ns) decays. The ultrafast decays are attributed to fast internal conversion between the first

two excited singlet states (1La and 1Lb) of Trp, the intermediate decays to local wobbling

motions of Trp, and the slow decays to tumbling motions of the proteins (28, 37). Similar

values for the wobbling motions (φTrp) and their cone semiangles (θ) were observed for

each series of S31L and A37L proteins (table 3.1); see material and methods for details.

Both mutation of residues around Trp and fluorination of the protein hydrophobic

core can affect the environment of the probe and change the protein structure and/or the

161 dynamic properties of the hydration layer. In many cases these properties are related to

one another. The minimal change in the steady-state fluorescence spectrum caused by

replacement of Leu by Tfl or Hil suggests similar features of the hydration region probed

by Trp (e.g., the effective number of water molecules in the hydration shell). In addition,

the similarity of the Trp wobbling angle of the Leu-, Tfl-, and Hil-forms of the proteins

suggests similar organization and flexibility of neighboring residues around the probe

(28). All these features make it possible to compare the dynamic properties of protein

hydration for most of the proteins in the A1m, S31L, and A37L proteins containing Leu,

Tfl, and Hil. For A1m-H, we note that the wobbling angle of Trp was found to be 33°,

which is significantly higher than the 17°–21° wobbling angles determined for all of the

other proteins. This result indicates that the organization of local residues or the

flexibility of the local Trp environment in A1m-H differs from that in the other proteins,

despite the lack of global structural changes observed by circular dichroism or

sedimentation velocity measurements. The abnormal behavior of the A1m-H variant is

also observed in the fluorescence lifetime measurements. Every protein except A1m-H

displayed a short-lifetime component of a few hundred picoseconds, present at all

wavelengths. These types of quenching processes have been attributed to Trp interactions

with nearby charged residues (38-40), and the absence of such a feature in A1m-H again

indicates that this protein has a local structure different from those of the other eight

proteins. The perturbation of local structure and, thus, local solvent exposure can result in

different hydration dynamics, making it unreliable to compare the dynamics of A1m-H to

those of the other A1m proteins. Accordingly, the dynamics obtained for A1m-H were

not used in the analysis that follows. Small shifts in the fluorescence emission maximum

162 and Trp wobbling angle were observed for A37L-L as compared to A37L-T and A37L-H

(see table 3.1 and figure 3.4). These differences may be significant enough to alter the

local environment surrounding the Trp probe, potentially complicating assignment of

changes observed in the hydration dynamics to a particular effect (e.g., changes in an

amino acid close to Trp). Despite these concerns, the dynamics results for A37L-L

remain consistent with the conclusions of the paper (see below).

Our stringent standards for comparison of hydration dynamics between modified

proteins require that there be (i) no global change in protein structure as measured by

circular dichroism spectroscopy and sedimentation velocity measurements; (ii) no change

in solvent exposure as measured by steady-state fluorescence maximum (± 1 nm); and

(iii) no change in local protein structure or flexibility as measured by fluorescence

anisotropy (± 1°). Seven of the nine proteins prepared in this study met all of these

criteria, and an additional protein, A37L-L, displayed changes just outside the margin of

error. Only one protein, A1m-H, showed changes significant enough to require us to

disregard the hydration measurements observed. Given the subtle effects of the chemical

environment on hydration dynamics, we will compare hydration results only within

protein families. Thus, our strongest conclusions will be drawn from observations made

on the S31L protein variants, and the data for A1m and A37L will be used as

corroborating evidence.

Ultrafast hydration dynamics. To investigate hydration dynamics at the protein

surfaces, we utilized a methodology developed by Zhong and coworkers for the

reconstruction of femtosecond-resolved fluorescence spectra (28, 41). As an example,

163 figure 3.6A shows several representative femtosecond-resolved fluorescence transients

recorded for A1m-T. The overall decay dynamics is retarded compared to that of free Trp

in buffer solution. Details of the results for all the protein samples are presented in table

3.1. The hydration dynamics of the proteins were well represented by triple-exponential

decays with distinctive timescales of 0.2–0.8, 1.4–6.1, and 10–61 picoseconds.

Relaxation occurring on a time scale of a few hundred femtoseconds to several

picoseconds is attributed to fast librational/rotational motions of bulk-type and local

water molecules around Trp. Observation of the fs component suggests that the Trp probe

in the test proteins is neither crowded by neighboring residues nor protected from

exposure to water (27, 28, 42). The slowest phase of hydration dynamics (on the

timescale of tens of picoseconds) is collective water network rearrangement coupled to

protein fluctuation dynamics (27, 42-44).

Several key features of the results (figure 3.7 and table 3.1) are summarized as

follows. First, S31L-T and A37L-T, in which Tfl lies close to Trp as well as in the

hydrophobic core, showed slower hydration dynamics than their Hil and Leu

counterparts, indicating that the fluorinated surface of the protein slows down the

hydration dynamics. For S31L-T, the timescales of local and collective water motions (τ2

and τ3, respectively) are increased by 2–5 times (3.0 and 48 ps) from those of S31L-H

(1.4 and 10 ps). The overall solvation of S31L-T is slower than that of S31L-L as well.

However, this difference is manifested as an increase in the contribution of the τ3

component to the overall solvation of Trp. The 70% increase of the relaxation energy

(333 cm−1) of the slowest component (E3) compared to that of the nonfluorinated S31L-L

(194 cm−1) is an indication of the dramatic slowing of hydration dynamics near the

164 fluorinated surface. For A37L-T, τ2 and τ3 are also retarded to a greater extent (3–5 times

slower) than those for A37L-H. These results suggest that replacement of Leu by Tfl

increases the residence time of water molecules near the Trp probe and/or the number of

water molecules influenced by the amino acid side chain.

Second, S31L-H and A37L-H showed similar or even faster hydration than their

Leu counterparts. This result indicates that in the comparison between hydrocarbon

residues, increasing the hydrophobic surface area results in faster motion of water

molecules around the residue. It should be noted that, for the S31L series, hydration is

greatly accelerated when Leu is replaced with Hil. This pronounced hydrophobic effect

(due to the increase in the size of the residue) on the hydration is counteracted by

fluorination of leucine, resulting in slowing dynamics for S31L-T. On the other hand, for

A37L, increasing the size of the hydrophobic surface does not appear to affect the

hydration dynamics as greatly. Therefore, the retardation of the dynamics upon

fluorination is much more pronounced for A37L-T than for the corresponding

fluorination of S31L. Finally, A1m-L and A1m-T, which differ from one another only in

the nature of their hydrophobic cores, exhibited almost identical hydration dynamics.

Fluorination of the hydrophobic core of a helix-bundle protein can affect protein

dynamics (45). However, the nearly identical hydration dynamics for A1m-L and A1m-T

spanning a few hundred picoseconds indicates that modification of the hydrophobic core

of A1m does not affect protein motions that are coupled to local hydration dynamics at

the surface of the protein on the timescales examined here (46).

165

Discussion

Protein surfaces are heterogeneous, consisting of polar, hydrophilic, and

hydrophobic residues, and it is intriguing to consider how the heterogeneous surface

chemistry affects the behavior of water molecules in the protein hydration layer. Head-

Gordon and coworkers have reported heterogeneous water dynamics in the first hydration

shell of model peptides (N-acetyl-leucinemethylamide and N-acetyl-glycinemethylamide),

with faster water motions near the hydrophobic side chains than near the hydrophilic

backbone (47, 48). Similar results have been reported for molecular dynamics simulation

studies of a folded β-hairpin peptide (30). In addition, Qiu et al. showed that mutation of

charged or polar residues of the enzyme staphylococcal nuclease into more hydrophobic

residues (Ala), resulted in faster hydration dynamics; this result was attributed to the lack of

strong interaction between the charges (or dipoles) of the mutated protein and the

surrounding water (27). This observation can be understood in that the elimination of

specific interactions between hydrophobic residues and water molecules causes a lower

number of hydrogen bonds between water and a hydrophobic surface compared to those

near a hydrophilic surface, thus allowing water molecules to reorient more freely.

Computational studies have suggested that water layers adjacent to extended hydrophobic

surfaces of low curvature are of lower density than those around hydrophilic and small

hydrophobic molecules, and are dynamic rather than static (49-56). X-ray reflectivity

experiments indicate submonolayer water depletion at hydrocarbon and fluorocarbon

surfaces (57). Our observation of accelerated hydration dynamics around larger Hil

residues as compared to the smaller Leu is consistent with these experimental and

computational results, supporting the idea that water molecules neighboring hydrophobic

166 side chains in the hydration layer of proteins are more dynamic than those around polar or

hydrophilic residues.

Even though a simple comparison suggests that Tfl should be more hydrophobic

than Leu by virtue of its larger surface area, introduction of a Tfl residue adjacent to the

Trp probe caused retardation of the local hydration dynamics, in contrast to the results

obtained when Leu was replaced with Hil (figure 3.7). These results suggest that hydration

dynamics around fluorinated amino acid side chains cannot be explained exclusively by the

increase in residue size. The C-F bond is assumed not to be involved in hydrogen bonding

with liquid water, largely because of its low polarizability (17). However, replacement of

Leu by Tfl introduces a strong dipole at the fluorinated protein surface. Lee and coworkers

have shown that introduction of CF3 groups reduces the contact angle of water on self-

assembled alkanethiol monolayers (23), an effect that they attribute to dipolar interactions.

Our results suggest that such dipolar interactions can also slow water motions at fluorinated

molecular surfaces.

Fluorinated compounds are more hydrophobic than hydrogenated compounds of

equal carbon number (4, 5, 17-21), and the increase in hydrophobic character of

fluorocarbons has been ascribed to their increased molecular size (18, 58). This

interpretation appears to be consistent with the observation that the melting temperature of

A1-Tfl is 13 °C higher than that of A1-Leu (11), while Tm for A1-Hil is increased by 17 °C

in comparison to A1-Leu. However, the results reported here clearly indicate that the

chemical nature of the protein surface dictates the dynamics of solvent-protein interactions,

and that size effects alone cannot explain the altered solvation dynamics observed at

fluorinated protein surfaces.

167

Conclusions

The results reported here show that fluorinated amino acids influence hydration

dynamics at protein surfaces in a manner quite different from their hydrogenated

counterparts. In general, water-protein interactions dictate many processes crucial to

protein function including folding, dynamic motions, interactions with other biomolecules,

and enzymatic catalysis (26). The slower timescales of hydration dynamics observed near

fluorinated residues in proteins suggest that some of the water-mediated processes

processes listed above may be changed upon fluorination. Tailoring the dynamics of

protein-water interactions by the introduction of fluorinated residues may yield proteins

with functional properties, such as binding, molecular recognition, or catalytic activities,

that cannot be achieved with the canonical amino acids. Understanding hydration dynamics

at fluorinated molecular surfaces is a critical step toward exploiting the properties of

fluorine in biological systems.

Materials and Methods

Summary of protein expression and characterization. A1 variants A1m, S31L, and

A37L were expressed in 2×YT medium (which contains Leu) to yield proteins A1m-L,

S31L-L, and A37L-L, respectively, in Leu-free M9 minimal medium (12.8 g/L

Na2HPO4·7H2O, 3 g/L KH2PO4, 0.5 g/L NaCl, 1 g/L NH4Cl) supplemented with 19

canonical amino acids plus Tfl to give A1m-T, S31L-T, and A37L-T, and in Leu-free M9

medium supplemented with 19 canonical amino acids plus Hil to give A1m-H, S31L-H,

and A37L-H. The proteins were purified under denaturing conditions and dialyzed against

10 mM acetate (pH 4)/100 mM NaCl. The extent of replacement of Leu by Tfl in A1m-T,

168 S31L-T, and A37L-T, was determined by amino acid analysis to be 90%–91%. Leu

replacement by Hil in A1m-H, S31L-H, and A37L-H was analyzed by liquid

chromatography tandem mass spectrometry and determined to be at least 90% (figure 3.8).

See below for further details of expression, purification, and incorporation analysis.

Summary of steady-state measurements. Circular dichroism spectra were recorded on an

Aviv 62DS spectropolarimeter (Lakewood, NJ). Absorption spectra were collected using a

Cary 500 UV-Vis spectrophotometer and a 0.05 mm path length cuvette. Steady-state

fluorescence emission spectra were measured using a FluoroMax-2 fluorimeter (ISA-

Spex).

Summary of time-resolved fluorescence measurements. The experimental apparatus for

time-resolved measurements are detailed below. All fluorescence spectra and transients

were obtained by the excitation of samples (~550 µM) at 295 nanometers. The lifetime

components were obtained by global analysis of fluorescence transients collected using a

time-correlated single photon counting spectrometer. All transients show additional

multiple-exponential decay (at the blue side) and rise (at the red side) with time constants

spanning from a few hundred femtoseconds to several tens of picoseconds. In order to

extract hydration dynamics precisely, we reconstructed apparent and lifetime-associated

time-resolved fluorescence spectra with eight or nine transients at different wavelengths

covering the blue and the red sides (figure 3.6B). By fitting these spectra to lognormal

functions, we traced the time-dependent apparent emission maxima (νs) and lifetime-

associated emission maxima (νl) as plotted in figure 3.6C. Using Δν(t) = νs(t) − νl(t), we

169 correlated the extracted time-dependent spectral shift, Δν(t), to the hydration energy

relaxation, ΔEs (figure 3.7).

Materials. All restriction enzymes were purchased from New England Biolabs (Beverly,

MA). D,L-5,5,5-trifluoroleucine (Tfl) was purchased from Oakwood Products (West

Columbia, SC). DNA oligomers were synthesized at Qiagen (Valencia, CA) or Integrated

DNA Technologies (Coralville, IA). (4S)-2-amino-4-methylhexanoic acid

(homoisoleucine, Hil) was prepared according to the methods of O’Donnell and Eckrich

(59) and Dorizon and coworkers (60).

Plasmid construction. An EcoRI/HindIII fragment of pQEA1 (11) containing the A1

coding sequence was ligated into EcoRI/HindIII-digested pQE-80L (Qiagen) to yield pQE-

80L/A1. The Asp residue at position 34 of A1 was mutated to Trp by site-directed

mutagenesis. The resulting plasmid was designated pQE-80L/A1m. A Leu codon was

introduced into either position 31 or position 37, yielding pQE-80L/S31L and pQE-

80L/A37L, respectively. Plasmid pA1EL (12), which encodes both the protein A1 and a

constitutively expressed leucyl-tRNA synthetase (LeuRS), was mutated using similar site-

directed mutagenesis techniques. A Trp codon was introduced first at position 34 of A1,

yielding pA1mEL. Introduction of leucine codons into either position 31 or position 37

resulted in the plasmids pS31LEL and pA37LEL, respectively.

Expression of fluorinated proteins. M9 medium supplemented with 0.4% glucose,

3.5 mg⁄L thiamine, 1 mM MgSO4, 0.1 mM CaCl2, 20 amino acids (40 mg⁄L), and 200 mg⁄L

170 ampicillin was inoculated 1:50 with an overnight culture (M9) of Escherichia coli strain

DH10B transformed with pQE-80L/A1m, pQE-80L/ S31L, or pQE-80L/A37L and grown

at 37 °C with shaking. After each culture reached OD600 = 0.9–1.0, the cells were harvested

by centrifugation (6000 × g, 4 °C, 6 min) and washed twice with cold 0.9% NaCl. The cell

pellets were resuspended in M9 medium containing 19 amino acids (no Leu) and 1 mM

Tfl. Protein expression was induced 10 min after the medium shift by addition of IPTG to a

final concentration of 1 mM. After 3 h, the cells were harvested by centrifugation (6000 ×

g, 4 °C, 10 min), and the cells were stored at −20 °C at least 12 h before purification. In the

case of the protein S31L-T, one sample was made using the procedure for the production of

proteins containing Hil described below.

Expression of proteins containing homoisoleucine. M9 medium supplemented with

0.4% glucose, 35 mg⁄L thiamine, 1 mM MgSO4, 0.1 mM CaCl2, 20 amino acids (40 mg⁄L),

200 mg⁄L ampicillin, and 35 mg⁄L kanamycin was inoculated 1:50 with an overnight

culture (M9) of E. coli strain LAM1000 transformed with pREP4 and pA1mEL, pS31LEL,

or pA37LEL at 37 °C with shaking. After each culture reached OD600 = 0.9–1.1, the cells

were harvested by centrifugation (5000 × g, 4 °C, 15 min) and washed three times with

cold 0.9% NaCl. The cell pellets were resuspended in M9 medium containing 19 amino

acids (no Leu) and 0.5 mM Hil. Protein expression was induced 15 min after the medium

shift by the addition of IPTG to a final concentration of 1 mM. Cells were harvested by

centrifugation (5000 × g, 4 °C, 15 min), resuspended in Qiagen buffer B (8M urea,

100 mM NaH2PO4, 10 mM TrisCl, pH 8.0) and sonicated for 10 min total process time

171 with a pulse duration of 5 s and a wait duration of 5 s. The sonicated lysates were frozen at

−80 °C for at least 12 h before proceeding with purification.

Expression of hydrogenated proteins. Rich (2×YT) medium was used instead of

supplemented M9 medium. When the culture reached OD600 = 0.9–1.0, IPTG was added to

a final concentration of 1 mM. After 3 h, the cells were harvested by centrifugation (6000 ×

g, 4 °C, 10 min), and the cells were stored at −20 °C at least 12 h before purification. In the

case of S31L-L, one sample was expressed using the cell strain LAM1000 containing

pREP4 and pS31LEL and harvested using the procedure used for the production of proteins

containing Hil.

Protein purification. N-terminally histidine-tagged A1 variants were purified under

denaturing conditions by affinity chromatography using Ni-NTA resin (Qiagen,

Chatsworth, CA) according to the manufacturer’s instructions. For proteins containing Hil

and for one batch each of S31L-L and S31L-T, the lysates in Qiagen buffer B were thawed,

sonicated, and then clarified using centrifugation (~75,000 × g, 25 °C, 10 min). Imidazole

was added to Qiagen buffer C (50 mM) and Qiagen buffer E (250 mM) in order to improve

purification efficiency. The purified protein solutions were dialyzed against 10 mM sodium

acetate (pH 4)⁄100 mM NaCl, and were concentrated by ultrafiltration (Amicon Ultra-15

devices, mwco: 10,000 or 3000, Millipore, Billerica, MA). The protein concentration was

determined as measured by the absorbance at 280 nanometers of solutions, assuming

extinction coefficients of 5500 M−1 cm−1 (61).

172 Amino acid analysis and sedimentation velocity analysis. Amino acid analysis of

fluorinated proteins was performed at the W. M. Keck Facility at Yale University (New

Haven, CT) on a Hitachi L-8900 amino acid analyzer (San Jose, CA) after hydrolysis at

115 °C in 70% formic acid. Sedimentation velocity analysis was performed at the National

Analytical Ultracentrifugation Facility at the University of Connecticut (Storrs, CT) by

using a Beckman XL-I Analytical Ultracentrifuge at 20 °C. The rotor was accelerated to

55,000 rpm, and interference scans were acquired at 1 min intervals for 7 h. The data were

analyzed by using the program Sedfit (62) to obtain normalized c(s) versus sedimentation

coefficient plots (figure 3.3).

Mass spectrometry. Liquid chromatography tandem mass spectrometry (LC/MS/MS) of

proteins containing Hil or Tfl was performed at the Caltech Protein and Peptide Mass

Analytical Laboratory. Trypsinized samples were subjected to liquid chromatography on an

Eksigent (Dublin, CA) NanoLC-2D using a 6 cm long, 100 µm diameter C18 column,

followed by MS/MS on an Applied Biosystems (Foster City, CA) QStar XL instrument.

Data were analyzed using Analyst QS software provided by Applied Biosystems. Hil or Tfl

incorporation levels were estimated using information contained within extracting ion

currents (XIC) of trypsin-digested protein samples. For a given sample, a peak

corresponding to a peptide globally substituted with the noncanonical amino acid and

coding for multiple leucines was identified, and the related peak corresponding to

replacement at a fraction of the leucine positions was also identified. Determination of the

ratio of the partially substituted to globally substituted peak areas allowed for the

estimation of amino acid incorporation levels assuming that leucines in the fragment were

173 replaced statistically. An example calculation is shown in the subsection “LC/MS/MS

amino acid incorporation estimates,” which can be found below.

Time-correlated single-photon counting (TCSPC). The protein samples were prepared at

55 µM concentration in 10 mM acetate (pH 4)⁄100 mM NaCl solution. The TCSPC

measurements were performed by using femtosecond pulses (<100 fs) from a Ti-sapphire

oscillator (Spectra-Physics, Mai Tai HP). Laser output, of which the repetition rate was

attenuated from 80 to 8 MHz utilizing a pulse picker (Spectra-Physics, Model 3980-5), was

tuned to 885 nanometers and frequency-tripled to 295 nanometers using a time-plate tripler

(Minioptic Technology, TP-2000B) for selective excitation of Trp. The UV beam,

vertically polarized using a half waveplate, was introduced to a sample chamber and

focused onto the sample cell. The residual frequency-doubled beam from the tripler was

directed to a photodiode to trigger a TCSPC system (PicoQuant GmbH, FluoTime 200).

Typically, the energy of the excitation pulse (attenuated) at the sample was ~10 picojoules.

In a right-angle geometry, the emitted fluorescence was collected at a magic angle (54.7°)

with respect to the vertically polarized excitation beam and focused into a MCP-PMT

(Hamamatsu, R3809U), which is attached to a double monochromator. The photomultiplier

tube signal was routed to a time-to-amplitude converter as a start signal followed by a

constant fractional discriminator (PicoQuant GmbH, SPC 630). To avoid possible

photobleaching and photodegradation, samples were kept stirring using a micro magnetic

stirrer. In this configuration, the instrument response has a full width at half maximum of

~30 ps. Multiexponential decays convoluted with instrumental response functions were

analyzed using the FluoFit software package (PicoQuant).

174 Femtosecond fluorescence upconversion. The protein samples were prepared at 550 µM

concentration in 10 mM acetate (pH 4)⁄100 mM NaCl solution. An amplified Ti-sapphire

laser system (Spectra-Physics, Hurricane X) was used, which produces ~110 fs pulses

centered at 805 nanometers (fundamental), with a 1 kHz repetition rate and a 0.8 millijoule

energy. The output beam was split into equal parts to generate the pump and the gate pulse

trains. For the pump, the fundamental light was used to pump an optical parametric

amplifier (Spectra-Physics, OPA-800C), the infrared idler output of which was sum-

frequency mixed with the residual fundamental in a 0.5 mm thick β-barium borate (BBO)

crystal (type I), recompressed with a prism pair, and frequency-doubled to provide the 295-

nanometers pulses in a 1.0 mm thick BBO crystal. The pump pulses were focused, with a

24 cm focal length lens, on the rotating circular cell (1 mm thickness) containing the

sample. Typically, the energy of the pump pulse (attenuated) at the sample was ~200 nJ. At

these energies, the fluorescence signals from samples were linearly dependent on the pump

energy. To check for sample degradation during experiments, fluorescence spectra were

periodically measured right after the rotating cell by using a fiber-optic-coupled

spectrometer (Acton Research, SpectraPro-300i) coupled to a charge-coupled device

(Princeton Instruments, SpectruMM-256HB) before and after the collection of averaged

transients for each sample. No difference between the spectra was observed.

The forward-scattered fluorescence from excited samples was collected and

focused by two off-axis parabolic mirrors into a 0.5 mm thick BBO crystal. Cutoff filters

were placed between the mirrors to reject scattered laser light and pass the desired

fluorescence wavelengths. The gate pulses, attenuated to 23 µJ⁄pulse, passed through a

computer-controlled optical delay line and were noncollinearly overlapped with the

175 fluorescence in the BBO crystal. After the crystal, the upconverted signal was separated

from the gate beam and the fluorescence by using an iris, and was focused on the entrance

slit of a 0.25 m double-grating monochromator (Jobin Yvon, DH10) equipped with a

photomultiplier tube at the exit slit. Upconversion efficiency was maximized by angle-

tuning of the BBO crystal. The upconverted fluorescence transients were taken at the magic

angle (54.7°) of the pump polarization relative to the gate polarization, parallel to the

acceptance axis of the upconversion crystal, in order to eliminate the influence of induced

sample anisotropy on the signal. The photomultiplier output was amplified (Stanford

Research Systems (SRS), SR445) and processed by a gated integrator (SRS, SR250). The

temporal response of the instrument was typically 350–450 fs. The observed fluorescence

transients were fit to theoretical functions, using a Scientist nonlinear least-squares fitting

program (Micromath), for the convolution of the Gaussian instrument response function

with a sum of exponentials. All experiments were carried out at an ambient temperature of

~24 °C, and all fluorescence transients were obtained by the excitation of samples at

295 nanometers.

For fluorescence anisotropy measurements, the pump-beam polarization was

rotated either parallel or perpendicular to the acceptance axis of the upconversion crystal to

collect the parallel (I||) and perpendicular (I⊥) signals, respectively. These transients were

used to construct time-resolved anisotropy: r(t) = (I|| – I⊥)/(I|| + 2I⊥). The results of the time-

resolved anisotropy are shown in figure 3.5. The ultrafast depolarization time constant, φI,

attributed to fast internal conversion between the first two excited singlet states (1La and

1Lb) of Trp, varies dramatically with the time resolution. This process has a timescale of

~100 fs (63, 64). The limited resolution of our current apparatus (350–450 fs) does not

176 allow us to fully resolve these dynamics and gives rise to a large uncertainty in the value of

φI. The variability of φI will impact the fit of φTrp. The uncertainty in the amplitude of the

anisotropy, rTrp and r∞, is not however affected by the limited time resolution of our data.

Therefore we use the wobbling cone angle to reveal details about the local crowding near

Trp. The wobbling cone angle is given by 1 – rTrp /(rTrp + r∞) = [(3cos2q – 1)/2]2 (37), and

only depends upon the amplitude of the tryptophan wobbling motion and the anisotropy

due to the rotation of the molecule.

LC/MS/MS amino acid incorporation estimates. LC/MS/MS was used to estimate the

replacement levels of leucine in some protein samples. Figure 3.8 depicts the total ion

currents (TIC, figure 3.8A) and three extracted ion currents (XICs, figure 3.8B–D) from a

digested A1m-H sample. The large peak in figure 3.8B corresponds to a peptide in which

all of the Leu residues are replaced by Hil, the smaller peak in figure 3.8C corresponds to a

mixture of two peptides containing one Leu and one Hil residue, and the very small peak in

figure 3.8D corresponds to a peptide containing only Leu residues. The areas in the three

XICs allow determination of the extent of incorporation of noncanonical amino acids in

place of leucine. Assuming that there is a probability p of homoisoleucine substitution in

place of leucine, the distribution of peak areas should correspond to the binomial

distribution,

€

A (1− p)2 + 2(1− p)p + p2[ ], (3.1)

177 where A is a multiplication factor equal to the total area of the three peaks and the three

terms of the polynomial correspond to nonsubstituted, singly substituted, and doubly

substituted peptides, respectively (the term for singly substituted peaks takes into account

both positional isomers of singly substituted peptides). Because these three terms represent

the only combinations of substitutions possible in the peptide, the relationship

(3.2)

also holds. The ratio between two peaks in a peptide series depends only on the probability

of incorporation and not on the value of A. Therefore, the ratio of two peaks from

experimental data can be used in order to get an estimate of p. The ratio of the peak areas of

singly substituted to doubly substituted peptides is

. (3.3)

Rearranging the above expression,

, (3.4)

where X is the experimentally observable ratio of singly substituted to doubly substituted

peptides. Solving for p gives

178 , (3.5)

or

, (3.6)

with the root of interest being the nonzero root. Substituting for the ratio of peak areas

gives an estimate of the incorporation level p.

In some cases, peaks corresponding to peptides containing three leucine or leucine

analogs were observed and used to quantify incorporation levels. In these cases, the peak

area distribution is represented by

, (3.7)

with

. (3.8).

Defining the ratio of doubly substituted to triply substituted peak area as X, substituting X

into equation (3.8), and solving for p yields the nonzero root

, (3.9)

179 again enabling an estimation of the incorporation level of noncanonical amino acids in

place of leucine. In some cases, the tandem mass spectrometry did not enable positive

identification of all possible positional isomers of a peptide. For example, in some cases,

only two out of three of the possible doubly substituted positional isomers containing three

possible substitution locations were identified in the tandem mass spectrometry data. In

these cases, X was multiplied by an appropriate factor to account for peptides that were not

observed (again invoking the assumption of completely random incorporation). Using the

above example, when only two out of three doubly substituted peptides could be identified,

X was multiplied by a factor of 1.5 in order to estimate what the peak area ratio would have

been with all three peaks present in equal weights. Using this methodology, the

homoisoleucine-containing proteins used were found to have 90% or greater Hil in place of

Leu, and the sample of S31L-T that was analyzed in this fashion contained approximately

99% Tfl in place of Leu. These results were obtained by examining three separate series of

peptides from each protein sample. These peptides had the following sequences:

AEIGDLNNTSGIR, GSHHHHHHGSMASGDLENEVAQLER, and SLEWEAAELEQK

(A1m), LLEWEAAELEQK (S31L), or SLEWEALELEQK (A37L).

Acknowledgements

We thank Professor Thomas Miller for helpful discussion, J. D. Fisk for synthesis

of homoisoleucine, and the referees for their thoughtful comments on the original

manuscript. This work is supported by National Institutes of Health (NIH) Grant

GM62523, National Science Foundation (NSF) Grant DMR-0964886, Office of Naval

180 Research (ONR) Grant N00014-03-1-0793, a Samsung Scholarship (to T. H. Y.), and a

National Defense Science and Engineering Graduate Fellowship (to J. A. V.).

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187

H2N COOH H2N COOHH2N COOH

CF3

1 2 3

Scheme 3.1. Amino acids used in study. 1, 5,5,5-trifluoroleucine (Tfl). 2, leucine (Leu). 3,

(4S)-2-amino-4-methylhexanoic acid (homoisoleucine, Hil).

188

Figure 3.1. Protein sequence and structure. A) Helical wheel diagram and B) amino acid

sequence of the A1 protein. The Asp residue at the f position of the third heptad (position

34) was replaced by Trp to yield a variant of A1 designated Alm. A Leu codon was

introduced at the c position of the third heptad (position 31, dark blue) or at the b position

of the fourth heptad (position 37, dark blue) to yield S31L and A37L, respectively. Side

views of C) A1m, D) S31L, and E) A37L.

189

Figure 3.2. Circular dichroism. Wavelength scans of A1m and variant proteins at 25 ºC.

The protein samples were prepared at 20 µM concentration in 10 mM acetate (pH 4)/100

mM NaCl solutions.

190

Figure 3.3. Normalized plots from the Sedfit c(s) analysis for A) A1m-L, B) S31L-L,

C) A37L-L, D) A1m-T, E) S31L-T, F) A37L-T, G) A1m-H, H) S31L-H, and I) A37L-H.

The protein samples were prepared at 550 µM concentration in 10 mM acetate

(pH 4)⁄100 mM NaCl solution. The broad, single peak of the A37L-L trace may be the

result of an equilibrium mixture between trimeric and higher-order species. It seems likely

that the breadth of the traces derived from sedimentation velocity analysis of A1m-T,

S31L-T, and A37L-T is a consequence of incomplete replacement of Leu with Tfl.

191

Figure 3.4. Steady-state UV-visible absorption (red) and fluorescence emission spectra

(black) of proteins excited at 295 nanometers. The protein samples were prepared at

550 µM concentration in 10 mM acetate (pH 4)⁄100 mM NaCl solution.

192

Figure 3.5. Time-resolved anisotropy, r(t), of the proteins. All anisotropy decays were

fitted to r(t) = rI exp(–t/fI) + rTrp exp(–t/fTrp) + r∞, where rI is the initial ultrafast anisotropy,

rTrp is the Trp motion-related anisotropy (value given in parentheses in each panel), r∞ is

the offset anisotropy, fI is the initial ultrafast internal-conversion time constant of Trp (≤1

ps), and fTrp is the Trp-rotational correlation time constant.

193

Figure 3.6. Hydration dynamics. Experimental determination of local hydration dynamics

at the surface of A1m-T, excited at 295 nm. A) Representative femtosecond-resolved

fluorescence up-conversion transients. B) Normalized time-resolved fluorescence spectra at

different time delays. The steady-state emission spectrum is also depicted (dotted line). C)

Time-dependent spectral shift of the apparent emission maxima (νs) and the lifetime-

associated (population) emission maxima (νl). Inset: entire evolution of νs and νl.

194

Figure 3.7. Hydration energy relaxation. Comparison of the hydration-correlated energy

relaxation, ΔEs(t), probed by Trp emission. Top panel: The solvation-energy relaxation data

for A1m proteins. Data for free Trp in the same buffer is also depicted for comparison.

Middle panel: Solvation energy relaxation data for S31L analogs. Bottom panel: solvation

energy relaxation data for A37L analogs. Insets: enlargement of the early-time hydration

behavior.

195

Figure 3.8. LC/MS/MS of trypsinized A1m-H. A) TIC of digested protein sample. B)–D)

XICs of peptides containing B) two, C) one, or D) no leucine to homoisoleucine

substitutions in the peptide SLEWEAAELEQK. The ratios of the peak areas obtained in

the XICs can be used to estimate the extent of leucine replacement in the protein sample.

Peptide masses: SXEWEAAEXEQK: 2+ ion: 730.86 Da observed, 730.87 Da expected.

SLEWEAAEXEQK, SXEWEAAEXEQK: 2+ ion: 723.88 Da observed, 723.86 Da

expected. SLEWEAAELEQK 2+ ion: 716.87 Da observed, 716.85 expected.

196 Table 3.1. Fluorescence emission maxima (λmax), hydration-correlated energy relaxation [ΔEs(t)], and depolarization dynamics [r(t)]

ΔEs(t)* r(t) † Sample λmax, nm

τ1, ps τ2, ps τ3, ps E1, cm−1 E2, cm−1 E3, cm−1 rTrp θ, °

Trp 353 0.30 1.5 13 883 682 18 0.196

A1m-L 352 0.30 2.1 31 610 646 171 0.056 21

A1m-T 352 0.28 2.5 31 877 568 161 0.049 20

A1m-H 352 1.9 34 0 2138 500 0.058 33

S31L-L 349 0.53 3.6 40 580 450 194 0.030 18

S31L-T 349 0.79 3.0 48 607 375 333 0.044 19

S31L-H 349 0.21 1.4 10 308 492 344 0.023 18

A37L-L 352 0.31 1.7 13 685 522 157 0.055 20

A37L-T 350 0.56 6.1 61 685 324 108 0.030 17

A37L-H 349 0.34 2.2 21 443 625 128 0.034 17

* All hydration-correlated energy relaxation dynamics were fitted to ΔEs(t) = E1exp(−t/τ1) + E2exp(−t/τ2) + E3exp(−t/τ3). † Refer to materials and methods for anisotropy analysis detail.